DEVELOPMENTAL

BIOLOGY

Variations

46, 430-435 (19%)

in the Amounts

III during

of RNA Polymerase

Preimplantation

Development

Forms I, II and

in the Mouse

LARRY R. VERSTEEGH, TIMOTHY F. HEARN AND CAROL M. WARNER’ Department

of Biochemist?

and Biophysics Accepted

Iowa State Uniaersity,

Ames, Iowa 50010

May 12, I975

DNA-dependent RNA polymerase has been measured at various stages of preimplantation development in mouse embryos. The total RNA polymerase activity per embryo increases rapidly from the 8-cell stage to the blastocyst stage. Studies with low cY-amanitin concentrations, which inhibit form II RNA polymerase, and high cu-amanitin concentrations, which inhibit both form II and III RNA polymerases indicate that the relative proportions of the three forms change significantly during preimplantation development. The changes which occur in the types and levels of RNA polymerase appear to parallel corresponding changes in the synthesis of the major classes of RNA. INTRODUCTION

Eukaryotic cell nuclei contain multiple forms of DNA-dependent RNA polymerase (1). The various enzyme forms appear to be involved in the specific transcription of different types of RNA. RNA polymerase I, which is localized in the nucleolus, synthesizes a ribosomal-type RNA, whereas polymerase II, which is found in the nucleoplasm, produces a DNA-like RNA (2, 3). RNA polymerase III, present in the nucleus and/or cytoplasm was recently shown to be involved in the synthesis of tRNA and 5S RNA (4). The existence of multiple forms of DNAdependent RNA polymerase, having different transcriptional specificities, suggests that this enzyme may play some role in the control of transcription. The question of whether or not the levels of RNA polymerase correspond to the levels of RNA synthesis in early embryos has not, at present, received a conclusive answer. The changes in rate of total RNA synthesis during sea urchin development have been shown to be correlated with the variation in levels of RNA polymerase activity (5). Furthermore, the variations in the relative amounts of form I and form II ’ To whom correspondence

should be addressed. 430

Copyright 0 1975 by Academic Press, Inc. All rights of reproduction in any form reserved.

polymerase seem to parallel changes in the types of RNA synthesized. Studies with Xenopus laeuis, however, failed to produce evidence of any correlation in RNA polymerase levels and the amounts or classes of RNA synthesized in early embryos (6, 7). In two previous studies (8,9) we reported the existence of three forms of DNAdependent RNA polymerase in adult mouse liver nuclei. Two of the forms, IA and IB, were insensitive to inhibition by cu-amanitin whereas the third form, form II, was found to be completely inhibited by low concentrations of the inhibitor. Form III RNA polymerase is now known to be inhibited by high concentrations of CYamanitin (4, 9). We have taken advantage of the selective action of varying concentrations of cr-amanitin to investigate the levels of the various forms of RNA polymerase during preimplantation development in the mouse. MATERIALS

AND

METHODS

The superovulation of mice, embryo collection, homogenization procedure, and RNA polymerase assay conditions have been reported previously (10). The final reaction mixture (75 ~1) contained: 36 pg calf thymus DNA, 50 mM Tris-HCl, pH 7.9, 2 mM MnCl,, 1.6 mM MgCL, 6 mM

131

BRIEFNOTES

NaF, 8 mM KCl. 0.5 mg/ml BSA, 0.3 mM spermine, 0.5 mM dithiothreitol, 0.6 mM ATP, 0.6 mM GTP, 0.6 mM CTP, 0.004 mM 3H-UTP (New England Nuclear, specific activity 26.9 Ci/mmol), 3 pg pyruvate kinase (Sigma, 320 units/mg), 1 mM phosphoenolpyruvate, 0.09 M tNH,),SO,, 8.3$ glycerol. 0.03 mM EDTA. The reaction mixture was incubated at 37” for 10 min and the amount of incorporated 3H-UMP determined by the method of Litman (11). Incorporation of labeled precursor is linear for at least 1 hr. Assays for RNA polymerase activity at 2-cell, g-cell, morula, and blastocyst stages were conducted with 600. 200. 150. and 100 embryos respectively. All assays were performed at least in duplicate. We have determined the error in the RNA polymerase activity assays in embryos to be * 120;. In a typical assay of 100 blastocysts. we measure -2000 cpm with a background of < 200 cpm. Assays for RNA synthesis in cultured embryos were carried out in 50 ~1 droplets of culture medium (12). which contained (27.8 Ci/mmole. New 18 PM [3H]uridine England Nuclear 1. The labeling period was for 5 hr for all cell stages. Incorporation of label into high molecular weight product is linear for the 5 hr labeling period for all cell stages studied (unpublished observations). Assays for RNA synthesis at 2-cell, g-cell, morula. and blastocyst stages were conducted with 100, 25, 20. and 20 embryos respectively. All assays were performed at least in duplicate. We have determined the error in the RNA synthesis assays in embryos to be *77. Following incubation. embryos were washed three times in 5 ml of culture medium, effectively removing any extracellular [3H]uridine. After washing, embryos were picked up in 50 ~1 of a 0.01 M Tris, pH 7.4. 0.50; BSA solution. Twenty five microliters of carrier RNA (20 pg) were added and the contents immediately frozen. After three freeze-thawings. the entire contents were spotted on DEAE-cellulose paper discs and assayed according to

a modification of the method of Litman (11). Further details of the RNA synthesis assay will be reported later. RESULTS

RNA Pol>rmeraseActicit?, in Preimplantation Mouse Em bpos Figure 1 shows the variation in the levels of RNA polymerase activity at various stages of preimplantation mouse embryo development. RNA polymerase activity, although not detectable at the e-cell stage. was measurable at all stages studied. The detection of enzyme activity is completely dependent on exogenous DNA in the reaction mixture (9). A large increase in enzyme activity is observed at about the g-cell stage. Figure 1 also shows the inhibition of RNA polymerase activity by 1.l pglml Lu-amanitin at 0.09 M ammonium sulfate, at the various stages of preimplantation mouse development. The inhibition by cu-amanitin decreases from 804 at 67 hr post-HCG (human chorionic gonadotropin) to 35[T( at the blastocyst stage (91 hr post-HCG).

RN.4 Swthesis in Preimplantation Embqos

Mouse

A second set of experiments, designed to measure RNA synthesis in preimplantation embryos in culture. was carried out on embryos at various stages of development. The results are shown in Fig. 2. RNA synthesis was detectable at all cell stages, including 2-cell embFos. The RNA synthetic activity at the 2-cell stage is at a comparatively low level and there is a large increase in RNA synthesis at about the g-cell stage which parallels the increase in RNA polymerase activity described above (see Fig. 1).

Relatice Amounts of RNA Pol>rmerase Forms I, II and III in Preimplantation Mouse Embryos To determine the relative amounts of form I, II, and III RNA polymerase activity

432

DEVELOPMENTALBIOLOGY

VOLUME 46. 1975

A concentration of 283 pg/ml inhibits form II activity, and in addition inhibits form III activity, leaving only form I uninhibited. It should also be noted that purified enzyme forms I and III are optimally active at low ionic strength, whereas purified form II enzyme is optimally active at high ionic strength. DISCUSSION

FIG 1. Activity of embryonic RNA polymerase as a function of time of development (04). % inhibition of embryonic RNA polymerase activity in the presence of 1.1 pg/ml cu-amanitin as a function of time of development (04). All assays were performed at least in duplicate at an ammonium sulfate concentration of 0.09 M. Two-cell, 8cell. morula and blastocyst embryos correspond to 42 hr. 66 hr. 82 hr. and 91 hr post-HCG, respectively.

It has been hypothesized that multiple forms of RNA polymerase may play some role in the control of early embryonic development (5, 13). If this is true, then one would expect to see some correlation of the amounts and types of RNA polymerases present with the amounts and types of RNA synthesized by the embryos. In this paper, we present the results of the first study of this type on a mammalian embryonic system. We have found that the total RNA polymerase activity per preimplantation mouse embryo increases nearly fivefold from the g-cell stage to the blastocyst stage of development (see Fig. 1). At the same TABLE

1

THE EFFECT OF WAMANITIN ON PREIMPLANTATION MOUSE EMBRYOS AT Low AND HIGH IONIC STRENGTH Time after KG

(hours)

Frc. 2. Embryonic RNA synthesis as a function of time of development. Details of the assay procedure can be found in “Materials and Methods.” All assays were performed at least in duplicate. It should be noted that the data is reported as SH-uridine incorporated, and not as total uridine incorporated into RNA, since the specific activities of the uridine pools in the embryos is unknown. Two-cell, S-cell, morula and blastocyst embryos correspond to 42 hr. 66 hr. 82 hr. and 91 hr postHCG, respectively.

Hours post-HCG

70.5 (S-cell)

79.5 (morula)

present during the various stages of preimplantation development, the embryos were 89.5 (blastocyst) assayed at high and low ammonium sulfate in the presence of 1.1 or 283 pg/ml CYamanitin. The results are presented in Adult liver cells Table 1. It is currently believed (4, 9) that 1.1 pg/ml of cr-amanitin completely inhibits form II enzyme activity, but has no effect on form I or form III enzyme activity.

cNSHo.,2’ Con&itration um 0.04 0.04 0.09 0.09 0.04 0.04 0.09 0.09 0.04 0.04 0.09 0.09 0.04 0.04 0.09 0.09

a-amanitin Concerttration (pg/ml) 1.1 283 1.1 283 1.1 283 1.1 283 1.1 283 1.1 283 1.1 283 1.1 283

% Inhibition of total enzyme activity/ embryo 22 23 76 76 20 25 68 69 10 24 34 36 11 25 30 35

BRIEF

time, RNA synthesis in cultured embryos shows an “apparent” 30-fold increase (see Fig. 2). The data in Fig. 2 correlates well with earlier studies in which only low levels of RNA synthesis were detected at the 2-cell stage, followed by a large burst of RNA synthetic activity at the 8-cell stage of development (14-25). Interpretation of experiments of this kind to give an “actual” rather than an “apparent” value for RNA synthesis is not possible at this time. One must ideally take into account differential precursor uptake, pool sizes, specific activity of the labeled precursor, and the stability of newly synthesized RNA. Although several studies (23, 24) have attempted to answer some of these problems, none have been completely successful. For instance, Daentl and Epstein (23) have reported that conversion of uridine to UTP is not a limiting factor in measuring RNA synthesis, using 3H-uridine as the precursor molecule. However, the specific activities of the precursor pools have not yet been determined in mouse embryos at any preimplantation stages (24, 26). We have chosen conditions for this study (5 hr labeling period, 18 PM 3H-uridine) which have minimized some of these concerns. The details of these studies will be published later. Our inability to measure RNA polymerase activity at the 2-cell stage is most likely due to limitations of the assay since we are able to measure RNA synthesis in cultured 2-cell embryos (see Figure 2). Also, (Yamanitin inhibits the first cleavage in uiuo indicating that RNA synthesis is necessary for cleavage very early in mammalian development ( 14). It seemed possible that RNase activity or the concentration of some enzyme inhibitor might be high at the 2-cell stage resulting in an apparent lack of RNA polymerase activity. To examine these possibilities two experiments were performed. First, one hundred 2-cell stage embryos were added to an amount of form II enzyme activity similar to that detected in the homogenate

NOTES

433

of 100 blastocysts. No inhibition of enzyme activity was observed. Second, RNA polymerase enzyme activity in 2-cell embryos was not detectable even in the presence of a well known RNase inhibitor (27). Therefore, absence of a high molecular weight RNA product in the RNA polymerase enzyme assay is not due to rapid degradation of the product by RNase. Thus, the lack of RNA polymerase activity at the 2-cell stage is probably the result of low levels of RNA polymerase and not high levels of RNase or some other enzyme inhibitor. Using cell number per embryo, as determined by Olds et al. (28). we have calculated that there is no significant change in the total RNA polymerase level per cell from the &cell stage (0.54 attomoles UMP incorporat.ed per minute per cell) to the blastocyst stage (0.60 attomoles UMP incorporated per minute per cell). However, the total “apparent” amount of RNA synthesized per cell increases 8.4 times from the 8-cell stage to the blastocyst stage. Even taking into account, as best as we can, differences in 3H-uridine uptake at the different stages of development, the “apparent” amount of RNA synthesized per cell increases (unpublished observations). This perhaps suggests that since the total amount of RNA polymerase per cell is constant, the total amount of RNA synthesized in preimplantation mouse embryos is subject to other, as yet unidentified, control processes. However, any calculations of either RNA synthesis or RNA polymerase activity per cell must be regarded with some caution. Experimentally we can only determine activities per embryo. For instance, in blastocysts, a certain total RNA polymerase activity may reflect an equal distribution of enzyme activity among all cells. Alternatively, a larger amount of enzyme activity may be found in the inner cell mass with a concomitant small amount of enzyme activity in the trophoblast cells, or vice versa. Thus, for the time being, the most useful comparisons are probably those made on a per embryo basis.

434

DEVELOP~IENTALBIOLOGY

As is shown in Fig. 1, there is a signif’icant change in the inhibition, by 1.1 pg/ml a-amanitin. of embryonic RNA polymerase activity from the g-cell to the blastocyst stage of development. Since the total RNA polymerase activity per cell does not change over this time period, the change in inhibition by 1.1 pg/ml a-amanitin suggests that the levels of the uninhibited enzyme forms (I and III), are increasing during preimplantation development, whereas the a-amanitin sensitive form of the enzyme (form II) is decreasing. Although a quantitative assessment of the relative amounts of the form I, II and III enzymes is not possible at this time, the data in Table I can be used in calculating qualitative changes in the various enzyme activities in whole embryos. First of all, from the data at 0.09 Mammonium sulfate and 1.1 pg/ml a-amanitin. it appears that the activity of form II decreases (by more than 50%‘) from the g-cell to the blastocyst stage (relative to the total activity at each stagei. From this, it follows that form (I plus III) activity is increasing during this same time. Furthermore, comparison of the data at 0.04 M ammonium sulfate and either 1.1 pup/ml or 283 pg/ml a-amanitin suggests that form III activity increases from the g-cell to the blastocyst stage of development. It is interesting to note that the inhibition data suggests that at the blastocyst stage the relative levels of the various enzyme forms is comparable to the levels observed in adult liver cells. These findings correlate nicely with the autoradiographic studies of Mintz (15) in which the RNA synthesized up to the 4-cell stage was located in the nucleoplasm and the RNA synthesized from the g-cell stage on was mostly nucleolar. Ellem and Gwatkin have shown the latter to be predominantly ribosomal RNA (18). We are currently investigating the effects of cu-amanitin on the types of RNA synthesized by preimplantation mouse embryos. In summary, during preimplantation development in the mouse, “apparent” in-

VOLUME 46, 1975

creases in total RNA synthesis are accompanied by a corresponding increase in the levels of total RNA polgmerase activity. Furthermore, increases or decreases in the types and amounts of the major classes ok cellular RNA appear to be paralleled b> increases or decreases in the corresponding class of RNA polymerase. This work was supported by a grant from the Population Council. New York, and in part by NIH Grant No. AI 11752-01 IMB. REFERENCES 1. RUTTER, W. J.. GOLDBERG, M. I.. and PERRIARD. J. C. (1974). “Biochemistry of Cell Differentiation.” (J. Paul. ed.), pp. 267-300. Universit! Park Press, Baltimore, Manland. 2. ROEDER. R. G., and RULER, W. J. (1970). Proc. Natl. Acad. Sci. 1r.S.A. 65, 675-682. 3. BLA~I, S. P.. INGLES. C. J.. LINDELL. T. J.. MORRIS. P. W.. WEAVER, R. F.. WEINBERG, F.. and RUTTER. W. d., (19701. Cold Spring Harbor S.ymp. Quant. Bid. 35, 619-657. 4. WEINMANN, R.. and ROEDER, R. G. (19X). Proc. Nat. Acad. Sri. L’.S.A. 71, 1790-179-i. 5. ROEDER. R. G., and RUTTER, W. J. (19701. Biochemistry 9, 2513-2553. 6. ROEDER. R. G.. REEDER, R. H.. and BROWN, D. D. (19701. Cold Spring Harbor Symp. Quant. Biol. 35,727-735. 7. ROEDER. R. G. (19741. J. Biol. Chem. 239, 219-256. 8. VERSTEECH. L. R., and WARNER, C. (19731. Biothem. Biophys. Res. Commun. 53,838-8&I. 9. VERSTEEGH, L. R.. and WARNER. C. M. (19751. Arch. Biochem. Biophgs. 168, 13:3- 114. 10. WARNER, C. M., and VERSTEEGH. L. R. (1974). Nature (London 1 248, 678-680. 11. Lrrh1.m. R. M. ( 1968t. J. Biol. Chem. 243, 6222-6233. 12. WHITTEN. W., and BIC~;ERS. J. (1968). J. Reprod. Fert. 17, 399-101. 13. CHAMBON, P., GISSENGER.F., KEDINGER. C., MANDEL, J. L., MEILHAC. M., and NLIRET, P. (1972). Karolinska Symp. Res. Methods Reprod. Endocrinol. 5 22’1-246. 14. GOLBUS, M. S., ~ALARCO, P. G., and EPSTEIN. C. J. (1973): J. Exp. Zoo/. 186, 207-216. 15. MINTZ. B. (1961). J. Exp. Zool. 157, 85-100. 16. THOMSON, J. L.. and BIGGER~, J. D. (1966). Exp. Cell Res. 41, 311-427. 17. MONESI, V.. and SALFI, V.. (1967) Exp. Cell Res. 46,632-635. 18. ELLEM, K. A. 0.. and GWATKIN. R. B. L. (1968). Develop. Biol. 18, 311-330. 19. WOODLAND, H. R.. and GRAHAhf. C. F. 11969).

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